Download PROCUSTE1 Encodes a Cellulose Synthase Required for Normal

The Plant Cell, Vol. 12, 2409–2423, December 2000, www.plantcell.org © 2000 American Society of Plant Physiologists
PROCUSTE1 Encodes a Cellulose Synthase Required for
Normal Cell Elongation Specifically in Roots and Dark-Grown
Hypocotyls of Arabidopsis
Mathilde Fagard,a,1 Thierry Desnos,a,2 Thierry Desprez,a Florence Goubet,a,3 Guislaine Refregier,a
Gregory Mouille,a Maureen McCann,b Catherine Rayon,a,4 Samantha Vernhettes,a and Herman Höftea,5
a Laboratoire
b Department
de Biologie Cellulaire, Institut National de la Recherche Agronomique, 78026 Versailles Cedex, France
of Cell Biology, John Innes Centre, Norwich Research Park, Norwich NR4 7UH, United Kingdom
Mutants at the PROCUSTE1 (PRC1) locus show decreased cell elongation, specifically in roots and dark-grown hypocotyls. Cell elongation defects are correlated with a cellulose deficiency and the presence of gapped walls. Map-based
cloning of PRC1 reveals that it encodes a member ( CesA6) of the cellulose synthase catalytic subunit family, of which
at least nine other members exist in Arabidopsis. Mutations in another family member, RSW1 (CesA1), cause similar
cell wall defects in all cell types, including those in hypocotyls and roots, suggesting that cellulose synthesis in these
organs requires the coordinated expression of at least two distinct cellulose synthase isoforms.
INTRODUCTION
Cellulose, the most abundant biopolymer, is produced by
plants, bacteria, and certain animal species; it consists of
simple linear (1→4)-␤–linked glucan chains assembled in
parallel arrays in semicrystalline microfibrils (Delmer, 1999).
In primary walls of most cotyledonous species, cellulose microfibrils constitute an extensible network cross-linked by
hydrogen-bonded xyloglucans embedded in a hydrophilic pectin matrix (Carpita and Gibeaut, 1993; McCann and Roberts,
1994). In growing cells, cellulose microfibrils are deposited
transversely to the elongation axis and are thus thought to
constrain growth in one direction. Cortical microtubule arrays often parallel the orientation of the microfibrils and appear to play a key role in controlling the orientation of the
microfibrils in the primary wall (Giddings and Staehelin, 1991;
Wymer and Lloyd, 1996). Not only do microfibrils control cell
shape, but also their orientation in meristematic tissues controls the formation of organ primordia (Green, 1994) and
thus plays a key role in the control of plant architecture.
1 Current address: Friedrich Miescher Institut, P.O. Box 2543, 4002
Basel, Switzerland.
2 Current address: Département d’Ecophysiologie Végétale et de Microbiologie/Laboratoire du Métabolisme Carboné, Commissariat à
l’Energie Atomique de Cadarache, 13108 St. Paul-lez-Durance Cedex, France.
3 Current address: Department of Biochemistry, University of Cambridge, Cambridge CB2 1QW, UK.
4 Current address: Department of Botany and Plant Pathology, Purdue University, West Lafayette, IN 47907-1155.
5 To whom correspondence should be addressed. E-mail hofte@
versailles.inra.fr; fax 33-1-30-83-30-99.
When cell expansion is arrested, a secondary wall is deposited within the bounds of the primary wall. The matrix
polysaccharide composition of the secondary wall differs
from that of the primary wall and frequently contains a high
proportion of lignin. In addition, microfibrils are mostly deposited in the secondary wall in helical arrays in successive
layers of alternating pitch (Vian and Reis, 1991). As a result,
this wall is much more rigid and provides strength and resistance against compressive forces, in contrast to the flexible
primary walls of organs, which maintain their shape as a result of turgor pressure within their tissues. In summary,
these observations show that a thorough understanding of
the molecular mechanism and the regulation of cellulose
synthesis is indispensable for understanding multiple aspects of plant development.
To date, cellulose synthesis remains poorly understood
(Kawagoe and Delmer, 1997; Delmer, 1999). UDP-glucose is
the direct substrate added to the nonreducing end of the
growing (1→4)-␤-glucan chain. In plants, a membranebound sucrose synthase, the enzyme that converts sucrose
to UDP-glucose, may be linked physically to the cellulose
synthase complex and may play an important role in channeling sucrose into the cellulose synthase complex (Amor et
al., 1995). The biochemistry of cellulose synthesis is extremely complex, and thus far no efficient procedure to assay this activity in vitro has been described (Delmer, 1999).
In contrast to other cell wall polysaccharides, which are synthesized in the Golgi apparatus and transported to the cell
surface by vesicles, cellulose is thought to be synthesized
by plasma membrane–bound complexes. These complexes,
which can be visualized in higher plants as hexameric
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clusters of particles on freeze-fractured plasma membranes,
have been named rosette terminal complexes (TCs) or simply rosettes (Emons, 1985).
Genes encoding homologs of the catalytic subunit of cellulose synthase in bacteria have been identified in higher
plants (Pear et al., 1996). The roles of at least two family
members (CesA1 and CesA7) in the synthesis of cellulose
were further demonstrated by the identification of mutants
in Arabidopsis. rsw1 carries a mutation in CesA1 that
causes a temperature-sensitive radial cell expansion defect
and a cellulose deficiency in all cell types investigated (Arioli
et al., 1998). In this mutant, moreover, the TCs disappear on
images of freeze-fractured plasma membranes. irx3 is mutated in CesA7 and shows a collapsed xylem phenotype
and a greatly decreased cellulose content specifically in
secondary cell walls (Taylor et al., 1999). These genes thus
appear to encode functionally specialized isoforms required
for cellulose synthesis in primary or secondary walls. Recently, Kimura et al. (1999) showed that antibodies generated against a member of the cellulose synthase catalytic
subunit family preferentially labeled TCs in freeze-fractured
plasma membranes of Vigna angularis; thus, TCs were definitively identified as the site of cellulose synthesis.
Besides the polymerization of (1→4)-␤-glucan, the assembly of the newly synthesized polymer into microfibrils
remains poorly understood. The analysis of rsw1 suggested
that the synthesis and formation of microfibrils are two independent processes. Indeed, at the restrictive temperature
and in the absence of functional rosettes, an ammonium oxalate–soluble, presumably noncrystalline, (1 →4)-␤-glucan
accumulated (Arioli et al., 1998). Similarly, cotton fibers
grown in vitro in the presence of the herbicide Novartis CGA
325⬘615, which also inhibits the formation of rosettes, produced (1→4)-␤-glucan, but also produced it in an alkali or
ammonium oxalate–extractable form (Delmer, 1999). These
observations are consistent with the view that RSW1 and related proteins polymerize (1→4)-␤-glucans but mediate
crystallization into microfibrils only when they are associated in TCs.
Cellulose synthase catalytic subunits are encoded by a
large gene family. In Arabidopsis, at least 10 isoforms are
recognized (http://cellwall.stanford.edu/cellwall and http://
www-plb.ucdavis.edu/labs/delmer/genes.html). The strong
mutant phenotypes for RSW1 (CesA1) and IRX3 (CesA7)
suggest a functional specialization for these isoforms. The
functions of the other family members remain to be determined. Besides the catalytic subunits, presumably other
genes are involved in cellulose synthesis. In the bacteria Acetobacter xylinum and Agrobacterium tumefaciens , several
genes besides the cellulose synthase catalytic subunit are
essential for cellulose synthesis, although their exact functions remain to be determined (Saxena et al., 1990; Matthysse
et al., 1995a, 1995b). Interestingly, one of the essential
genes in Agrobacterium encodes an endo-(1 →4)-␤-glucanase, which has been proposed to play a controversial role
in the cleavage of lipid-linked intermediates of the biosyn-
thetic pathway. In plants, a membrane-bound endo-(1 →4)␤-glucanase, KOR, also may play a role in cellulose synthesis, as suggested by the cellulose deficiency of kor mutants
(Nicol et al., 1998; M. Fagard, G. Mouille, F. Goubet, and H.
Höfte, unpublished data).
In this article, further insight is provided regarding the cellulose synthesis machinery of plants. We report the cloning of
PROCUSTE1 (PRC1), which was identified previously by mutations causing a growth defect in roots and dark-grown hypocotyls (Desnos et al., 1996). Here, we show that the
decreased cell expansion in the dark-grown hypocotyl is correlated with a deficiency in cellulose and the presence of gaps
in internal cortical and epidermal cell walls. PRC1 was cloned
and found to encode a novel member of the CesA family,
CesA6. Interestingly, both rsw1 and prc1 mutants caused a
cell elongation defect and cellulose deficiency in roots and
dark-grown hypocotyls; however, prc1 affected only a subset
of the cell types that were affected in rsw1. This suggests that
cellulose synthesis in this subset of cell types requires the coordinated activity of at least two CesA isoforms.
RESULTS
Identification of Additional prc1 Alleles
Fourteen new alleles of the prc1 locus were identified from
Arabidopsis populations mutagenized with ethyl methanesulfonate, T-DNA, or x-rays. Among them were the previously described quill (qui) mutants (Hauser et al., 1995),
which were shown to be allelic to prc1 (data not shown; see
below). qui1, qui2, and qui3 were renamed prc1-9, prc1-10,
and prc1-11, respectively. Figure 1A shows the hypocotyl
lengths of seedlings for nine prc1 alleles grown for 7 days in
complete darkness. Consistent with previous results (Desnos
et al., 1996), all alleles (regardless of ecotype) showed a
four- to fivefold decrease in hypocotyl length compared with
that of wild-type seedlings, whereas no large variations in
allele strength were observed (Figure 1A). Root length was
also determined for seedlings grown for 7 days in white light
on agar medium containing 4.5% sucrose (Figure 1B), conditions under which roots of qui alleles exhibit a strong
radial expansion phenotype (Hauser et al., 1995). As expected, prc1 roots were 1.5- to twofold shorter than were
wild-type roots, with greater variations between alleles than
those observed for dark-grown hypocotyls. On the basis of
these root lengths, prc1-3 and prc1-4 appeared to be the
strongest alleles.
Alterations in Cell Wall Structure in Dark-Grown
prc1 Hypocotyls
The prc1 phenotype can be phenocopied by treating wildtype seedlings with cellulose biosynthesis inhibitors such as
PRC1 Encodes a Cellulose Synthase
Figure 1. prc1 Mutants Show Reduced Elongation of Roots and
Dark-Grown Hypocotyls.
(A) Dark-grown hypocotyl lengths of wild-type (ecotypes Columbia
[Col-0] and Wassilewskija [WS]) seedlings and 11 prc1 alleles. Seedlings were grown for 7 days in total darkness on sucrose-free agar
medium.
(B) Root lengths of wild-type seedlings and 10 prc1 alleles. Seedlings were grown for 7 days in 16-hr-light/8-hr-dark cycles on agar
medium containing 4.5% sucrose.
prc1-1 through prc1-9 and prc1-11 are Col-0 alleles; prc1-10 and
prc1-12 are Ws ecotype.
Error bars indicate ⫾SD.
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2B, top arrow). Incomplete walls were found in epidermal,
cortical, and endodermal cell layers. The protruding wall ends
were frequently found to be linked by a membranous structure not stained by Calcofluor but observable with Congo
Red (Figure 2C) or other stains. This structure presumably
corresponds to cell membranes surrounding polysaccharidic material. The study of two other alleles ( prc1-3 and
prc1-7; data not shown) confirmed the presence of such incomplete cell walls. In some sections (both transverse and
longitudinal), cells filled with granular Calcofluor- and Congo
Red–stained material were observed (Figure 2B, bottom arrow); these were never observed in the wild type, and the
exact nature of the material remains to be determined.
Incomplete cell walls, referred to as “stubs,” have been
described in various mutants that are defective in cytokinesis (Liu et al., 1995a; Assaad et al., 1996; Lukowitz et al.,
1996). Because cortical and most epidermal cells in Arabidopsis hypocotyls have an embryonic origin and do not divide during postembryonic development, we examined
sections through prc1-1 embryos (illustrated for a torpedostage embryo in Figure 2D). Cell wall stubs were never observed. The same was true for mature embryos, or even for
transverse sections through mature hypocotyls grown in the
light (data not shown). Together, these observations indicate that cytokinesis is unaffected in prc1 and that the observed gapped walls in dark-grown prc1 hypocotyls arise
during cell expansion.
Incomplete walls were also observed in cortical and epidermal walls in dark-grown prc1-1 hypocotyls by scanning
electron microscopy on freeze-fractured material or by
transmission electron microscopy, as shown in Figure 3; incomplete walls were observed as frequently as on sections
prepared for light microscopy. This finding strongly suggests that the discontinuous walls were present in living tissue and were not artifacts of the sample preparation.
Furthermore, transmission electron microscopy revealed the
presence of fibrillar material in the wall gaps (Figure 3B).
Mutations at the PRC1 Locus and Cellulose Deficiency
2,6-dichlorobenzonitrile (DCB) or isoxaben (data not shown).
To look for potential cell wall defects, we studied transverse
sections halfway through hypocotyls of 4-day-old darkgrown seedlings after staining with Calcofluor or Congo Red
(Figure 2). Calcofluor stains cellulose, callose, and other
nonsubstituted or weakly substituted ␤-glucans (Maeda and
Ishida, 1967), whereas Congo Red more generally stains reducing sugars. As shown in Figure 2, no differences in the
numbers of cells or tissue layers were observed. However,
similar to the DCB-treated seedlings, prc1-1 hypocotyls
were larger than were wild-type hypocotyls, with more radially expanded cells in epidermis, cortex, and endodermis.
Interestingly, in almost every cross-section of dark-grown
prc1-1 hypocotyls, incomplete cell walls were found (Figure
prc1 cell walls were analyzed by using Fourier transform infrared (FTIR) microspectroscopy. This technique allows
rapid analysis of the cell wall composition of areas of the tissue as small as one or a few cells (McCann et al., 1997).
FTIR absorption spectra of 4-day-old dark-grown hypocotyls of wild-type and two mutant alleles of prc1 were collected from a region of ⵑ60 ⫻ 40 ␮m midway along the
hypocotyl and avoiding the central stele. The spectra correspond to the absorption in the mid-IR region (between wave
numbers 600 and 4000 cm⫺1) of essentially longitudinal and
transverse cell walls of epidermal and cortical cells (two layers of epidermis and four layers of cortex cells), both of
which are cell types that are affected by the prc1 mutations.
The spectra (20 for each condition) were analyzed by principal components (PCs) analysis (Kemsley, 1998), a statistical
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Figure 2. Dark-Grown prc1 Hypocotyls Contain Incomplete Cell Walls.
(A) and (B) Calcofluor-stained hypocotyl cross-sections of 4-day-old
dark-grown wild-type (A) and prc1-1 (B) seedlings. The top arrow indicates an incomplete wall and the bottom arrow indicates an abnormally filled cell in a prc1-1 hypocotyl section.
(C) Congo Red–stained section of a prc1-1 hypocotyl cell in which
the incomplete cell wall stubs are connected by a thin structure (arrow).
(D) Calcofluor-stained section through a prc1-1 immature embryo.
No incomplete wall or cell filled with stained material was observed.
Bars in (A) and (B) ⫽ 100 ␮m; bar in (C) ⫽ 30 ␮m; bar in (D) ⫽ 60 ␮m.
method that reduces the dimensionality of the data from
⬎100 variates (one every 8 cm⫺1 for the region from 1800 to
850 cm⫺1) to only a few, the PCs. The PCs are ordered in
terms of decreasing variance. Each observation (spectrum)
has a corresponding set of PC scores describing the variance of that spectrum relative to the mean of the population
for each PC. The PC scores of the spectra can then be plotted against one another to reveal patterns or structures in
the data (Kemsley, 1998). One can then mathematically derive a “spectrum” (a PC loading) from a PC to identify the
molecular factors responsible for the separation of groups
of spectra (Kemsley, 1998). As shown in Figure 4A, the analysis showed that wild-type spectra can be separated from
prc1 mutant spectra by using a combination of two PC
scores. PC1 explained 72% of the variance. The loading for
PC1 (Figure 4B) showed characteristics of purified cellulose
in the fingerprint region (peaks at 1038, 1064, 1100, and
1162 cm⫺1, respectively) and of protein (peaks at 1650 and
1550 cm⫺1). Peaks at 1650 and 1550 cm⫺1 were negatively
correlated with the cellulose fingerprint peaks. Because the
PC scores of the spectra of prc1 seedlings were negative
relative to the mean (Figure 4A), the data suggest that the
cell walls of prc1 seedlings are relatively richer in protein
and poorer in cellulose than the cell walls of wild-type seedlings. These results show that the changes in prc1 cell walls
are complex but that a major deficiency in crystalline cellulose is a part of the phenotype.
To confirm the observed changes chemically, walls of
4-day-old dark-grown wild-type and prc1-1 seedlings were
isolated and fractionated, and the neutral sugar and uronic
acid contents were determined for each fraction. In both
wild-type and prc1-1 seedlings, ⵑ2% of the fresh weight
consisted of cell walls. Wall material was extracted consecutively with 0.1 and 4 M KOH, yielding two fractions that
were enriched for pectins (fraction 1) and hemicellulose
(fraction 2), respectively, as shown by the neutral sugar and
uronic acid contents (Figures 5C and 5D). The residual fraction contained 60% glucose (Figure 5E), which indicates an
enrichment for cellulose, because as shown below, in the
culture conditions used, neither wild-type nor prc1-1 accumulated significant levels of (1→3)-␤-glucan (callose). Also,
staining with iodine failed to detect starch in wild-type or
prc1-1 hypocotyls (data not shown). The presence of minor
amounts of xylosyl (5%), rhamnosyl (6%), uronic acid (10%),
galactosyl (10%), and arabinosyl (6%) residues indicates
that a proportion of the noncellulosic polysaccharides also
resisted the alkaline treatment, suggesting an intimate association of these polymers with the cellulose microfibrils.
Similar observations were reported for cell walls from Arabidopsis leaves (Zablackis et al., 1995).
Strikingly, in mutant walls, the proportions of the three
fractions were altered markedly (Figure 5A). Compared with
wild-type walls, prc1-1 walls showed a relative increase in
fraction 2, a relative decrease in the residual fraction, and a
very minor decrease in fraction 1. Interestingly, in prc1-1,
the decrease in the residual fraction was not accompanied
by a change in the proportions of neutral sugars and uronic
acid, suggesting that with less crystalline cellulose present
as a substrate, less xyloglucan and pectin resisted the extraction procedure. The neutral sugar and uronic acid contents also remained unchanged for fractions 1 and 2, except
for an important decrease of galactosyl residues in fraction
1. This may be the result of a reduction in the number of galactan side chains on the pectins.
To confirm the reduced cellulose production in prc1, we
measured the incorporation of 14C-glucose into the nitric/
acetic acid–insoluble cell wall fraction of actively growing
3-day-old seedlings (Updegraff, 1969). This fraction is 97%
(1→4)-␤–linked glucose (Peng et al., 2000; data not shown)
and is generally considered to be crystalline cellulose. Figure 5B shows that in the wild type, 30% of the label in the
cell wall was incorporated in the cellulosic fraction. This proportion was reduced to 17 and 19% in prc1-1 and prc1-4,
respectively.
In conclusion, chemical analysis of the cell walls of mutant seedlings confirms the deficiency in cellulose observed
by FTIR microspectroscopy. Other changes detected included a relative increase in hemicellulosic polysaccharides
and qualitative changes in pectin composition.
PRC1 Encodes a Cellulose Synthase
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The inhibition of cellulose by the cellulose synthesis inhibitor DCB in Arabidopsis seedlings causes an accumulation
of callose (Nickle and Meinke, 1998). Callose accumulation
was also observed in embryos of the cytokinesis-defective
mutant cyt1 (Nickle and Meinke, 1998). We investigated
whether also in prc1 the cellulose deficiency was accompanied by increased callose production. Staining hypocotyl
sections as well as permeabilized seedlings with aniline blue
or its fluorescent derivative (Stone et al., 1984) revealed a
slight accumulation of callose in DCB-treated seedlings. In
prc1-1 hypocotyl sections, we occasionally observed a
weak staining, however much weaker than that observed in
cyt1 (data not shown).
Map-Based Cloning of PRC1
The prc1-1 mutation was mapped previously to the bottom
of chromosome 5, 1.7 centimorgans (cM) distal to marker
m211, in a small mapping population (Desnos, 1996; Desnos
et al., 1996). New mapping populations were established by
using a cross between prc1-1 and Landsberg erecta (Ler) as
well as by crossing prc1-1 to a line (in the C24 ecotype) carrying a T-DNA insertion close to the LFY3 restriction fragment length polymorphism marker (Lister and Dean, 1993;
Van Lijsebettens et al., 1996). By use of restriction fragment
length polymorphism and cleaved amplified polymorphic
sequence (Konieczny and Ausubel, 1993) markers, the prc1-1
mutation was mapped to a region of ⵑ120 kb covered by
P1 clones (Liu et al., 1995b) MVP7 and MXK3. As shown in
Figure 6A, prc1-1 is positioned 0.1 cM distal to MUB3.p2B
and 1 cM proximal to 4B2-L/2F2-E12.
From the genomic sequence of the 120-kb region containing the prc1-1 mutation, seven candidate genes were
selected. Polymerase chain reaction products for each of
these genes were amplified from genomic DNA isolated
from several prc1 alleles, sequenced, and compared with
the relevant wild-type sequences. Six prc1 alleles showed
nucleotide substitutions in the sequence of a predicted
gene between positions 18,535 and 23,313 on P1 clone
MVP7 (GenBank accession number AB025637). All six mutations caused premature stop codons in the predicted coding region. Interestingly, despite their provenance from
independent M2 families, prc1-1 and prc1-3 showed mutations at identical positions, as did prc1-4 and prc1-5, albeit
at a different location (Figure 6). The allele prc1-19 generated by T-DNA mutagenesis contained a 28-bp deletion
Figure 3. prc1 Dark-Grown Hypocotyls Display Abnormal Cell Wall
Structures.
(A) to (C) Transmission electron micrographs of cross-sections from
dark-grown hypocotyls (4 days old) of prc1-1 mutant ([A] and [B]) and
wild-type (C) seedlings in the cortical cell layer. Arrow in (A) indicates
an abnormal wall portion shown at a higher magnification in (B).
(D) and (E) Scanning electron micrographs of prc1-1 dark-grown hy-
pocotyls. Arrowheads indicate protruding stubs of incomplete cell
walls in the epidermal cell layer.
co, cortex cell; cw, cell wall; cy, cytoplasm; ep, epidermal cell; va,
vacuole. Bar in (A) ⫽ 1 ␮m; bars in (B) and (C) ⫽ 0.2 ␮m; bar in (D) ⫽
10 ␮m; bar in (E) ⫽ 20 ␮m.
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reported slightly truncated cDNA (AraxCelA; GenBank accession number AF062485) (Wu et al., 1998). The gene contains 13 exons and encodes a protein of 1084 amino acids
with a predicted molecular mass of 123 kD. PRC1 is a member of a large, ancient family of ␤-glycosyl transferases
present in bacteria, plants, and animals. In higher plants, a
subfamily of highly conserved proteins can be distinguished, two members of which (RSW1 and IRX3) have
been shown to be essential for cellulose synthesis in Arabidopsis. This subfamily, referred to as CesA, probably corresponds to the family of catalytic subunits of cellulose
synthase and has at least 10 members in Arabidopsis. According to the classification proposed by Delmer (1999) (see
also http: //www-plb.ucdavis.edu/labs/delmer/genes.html),
PRC1 corresponds to CesA6. The predicted protein structure is shown in Figure 6B.
The PRC1 protein contains the U1, U2, U3, and U4 domains that are characteristic of all ␤-glycosyl transferases
and that form the catalytic domain. A protein fragment of a
cotton homolog (CelA1) containing these motifs was shown to
bind UDP-glucose in the absence of calcium ions (Kawagoe
and Delmer, 1997; Delmer, 1999). All CesA family members
from plants contain the so-called plant conserved region
and a region that is hypervariable (HVR) between family
members. Allele prc1-9 is predicted to produce a protein
truncated at the N-terminus of the first transmembrane domain. In the other mutants analyzed, the predicted truncated product contains only the first two transmembrane
domains and part of the catalytic domain. All mutants investigated therefore are expected to be null alleles.
Figure 4. prc1 Dark-Grown Hypocotyl Cell Walls Are Cellulose Deficient.
(A) FTIR analysis of 4-day-old dark-grown hypocotyls. PC analysis
was performed by using 20 FTIR spectra from wild-type and two different prc1 alleles. PC1 explained 72% of the variance between
spectra from wild-type alleles and spectra from prc1 alleles. Col0,
Columbia.
(B) The PC1 loading showed positive peaks characteristic of cellulose in the fingerprint region (1162, 1108, 1061, and 1038 cm⫺1, respectively), indicating that prc1 dark-grown hypocotyls were
deficient in cellulose relative to the wild type. Two other major peaks
(1652, and 1548 cm⫺1, respectively) appeared negatively in this PCloading. These peaks corresponded to amide groups of protein,
suggesting an enrichment for protein in prc1 walls relative to the
wild type.
(Figure 6), which may be a hallmark of an aborted T-DNA integration.
Isoform of the Cellulose Synthase Family Encoded
by PRC1
The genomic sequence of PRC1 perfectly matches five expressed sequence tag (EST) sequences and a previously
Two CesA Isoforms Required for Normal Cell Expansion
in Roots and Dark-Grown Hypocotyl Cells
An intriguing observation is that two mutations in members
of the same gene family caused very similar phenotypes in
the same cell types. Indeed, the leaky allele rsw1-10 (H.
Höfte, unpublished data) and the null allele prc1-8 both
showed comparable decreases in the length of dark-grown
hypocotyls, accompanied by increases in width (Figure 7) in
both epidermal and cortical cells (data not shown). FTIR microspectroscopy also indicated a cellulose deficiency in epidermal or cortical cells (or both) in both mutants (Figure 4
and data not shown). The rsw1-10 mutation was additive to
prc1-8, as shown by the further shortened hypocotyl length
of the double mutant (Figure 7).
One explanation for these observations is that both gene
products have comparable functions and contribute to a
pool of enzymes that becomes limiting in a mutant background for one or the other locus. However, that situation is
unlikely because both mutations are recessive and no striking phenotype was observed in plants that are transheterozygous for both mutations, whereas if the hypothesis
described above were true, then the phenotype should be
equivalent to that of a single homozygote. In addition, as
PRC1 Encodes a Cellulose Synthase
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shown in Figure 8, even for prc1-8 segregating in a homozygous rsw1-10 mutant background, no gene dosage effects
were observed, as shown by the strict 1:3 segregation of the
additive phenotype. Therefore, even in a situation in which
the RSW1 gene product is growth-limiting, both gene copies of PRC1, rather than a single copy, need to be inactivated to result in a further reduction of hypocotyl length.
This strongly suggests that the presence of each protein is
critical for normal hypocotyl elongation, rather than the size
of the combined pool of the CesA1 and CesA6 proteins.
Functional Specialization within the CesA Family
The strong mutant phenotypes for IRX3 (CesA7), RSW1
(CesA1), and PRC1 (CesA6) indicate that the other family
members cannot compensate for the absence of their respective gene products. In some cases, this may be simply
a result of cell type–specific expression patterns. However,
as shown above, this does not seem to be the case, at least
not for RSW1 and PRC1. Alternatively, the sequences of different family members may have diverged as adaptations to
their specialized cellular roles. To reveal such specialized
functions, we investigated the evolutionary relationships between different family members in Arabidopsis and cotton, a
species for which a large number of CesA ESTs are available in public databases. The sequences of 10 Arabidopsis
CesA family members were compared with those of three
full-length cotton cDNAs and two genes identified by ESTs
from different cotton libraries. Because the cotton ESTs corresponded to single-pass sequences close to the 3⬘ end of
the mRNA, we chose the amino acid sequence corresponding to the HVR, which is localized between amino acids 690
and 743 close to the C-terminus of PRC1, to construct a
similarity tree. The use of this hypervariable protein segment
also maximized the chances to distinguish among the different family members, which in many cases are extremely
conserved. A sixth, more divergent, cotton sequence (Gh
EST-AI727450), presumably corresponding to a cellulose
synthase–like gene (http://cellwall.stanford.edu/cellwall), was
used as an outlier in the bootstrap procedure. As shown in
Figure 9, amino acid sequences from the five cotton genes
did not cluster together but in each case clustered with one
or more genes from Arabidopsis. This suggests that the
Figure 5. Chemical Analysis Confirms Cellulose Deficiency in DarkGrown prc1 Seedlings.
(A) Walls were purified and extracted successively with 0.1 M KOH,
yielding a pectin-rich fraction (fraction 1), and 4 M KOH, yielding a
hemicellulose-rich fraction (fraction 2). The residual fraction was enriched for a glucan corresponding to cellulose (see text). prc1-1
walls were deficient for the residual fraction relative to the wild type,
indicating a deficiency in cellulose for this mutant. Col0, Columbia.
(B) Incorporation of 14C-glucose (14C) in the acid-resistant cellulosic
fraction in wild-type and prc1 alleles. Each value represents the
mean of five independent measurements. CW, cell wall.
(C) to (E) Sugar composition of fraction 1 (C), fraction 2 (D), and the
residue (E). Only fraction 1 reproducibly showed qualitative differences in sugar composition. r, rhamnose; f, fucose; a, arabinose; x,
xylose; m, mannose; ga, galactose; gl, glucose; u, uronic acid.
Error bars indicate ⫾SD.
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The Plant Cell
HVR is hypervariable among different family members within
a species but is conserved between the orthologs in different species. A very similar tree was obtained when we used
the complete amino acid sequences of the Arabidopsis
genes and the three full-length cotton genes (data not
shown). In conclusion, the divergence between the family
members, at least those represented by the five cotton
ESTs, must have predated the divergence between Brassicaceae and Malvaceae. Interestingly, RSW1 and PRC1 also
had their respective orthologs in cotton.
Expression Patterns of PRC1 and RSW1 mRNAs
RNA gel blot assays were performed with probes specific
for sequences corresponding to a variable region in the respective N termini of PRC1 and RSW1 (Figure 10). Both
PRC1 and RSW1 mRNA were detected in all organs investigated, except in mature flowers. Although the transcript
abundance of PRC1 was less than that of RSW1, both
mRNAs showed similar expression patterns. In developing
seedlings, RSW1 and PRC1 mRNAs were high initially and
decreased at later growth stages. Expression was high in
elongating stems but dropped in the stem sections in which
secondary wall synthesis takes place. Interestingly, despite
the light-dependent hypocotyl phenotype of prc1, we did
not obtain evidence for a regulation of PRC1 at the transcript level: indeed, transcript quantities did not differ between 4-day-old dark-grown and light-grown seedlings.
Similar results were obtained by comparing distributions
of the EST sequences corresponding to RSW1 and PRC1
obtained from tissue-specific EST libraries (http://cellwall.
stanford.edu/cellwall). For instance, in four EST libraries
separately covering etiolated seedlings, above-ground tissues,
roots, and siliques, we found, respectively, 1, 5, 5, and 1 EST
for PRC1 and 4, 13, 18, and 2 ESTs for RSW1. This confirms
that PRC1 expresses fewer transcripts than RSW1, that the
tissue distribution of each is similar, and that PRC1 mRNA is
present in both dark-grown and light-grown tissues.
Figure 6. The PRC1 Gene Is Isolated Through Map-Based Cloning.
(A) Physical map of the region of chromosome (chr.) 5 containing the
PRC1 gene. The prc1-1 mutation was mapped to the bottom of
chromosome 5, south of molecular marker LFY. New markers positioned the prc1-1 mutation between markers Mub3.p2B and 2F2E12. The number of recombinant chromosomes is indicated in parentheses. Sequence analysis of candidate genes between these
two markers showed that six prc1 mutant alleles carried a point mutation or a deletion in the CesA6 gene carried by P1 clone MVP7.
The PRC1 (CesA6) gene comprises 13 exons (filled boxes) and 12
introns. The prc1 mutations that were identified are indicated.
(B) Predicted topology of the PRC1 protein. The protein is predicted
to contain eight membrane-spanning helices (indicated by the eight
gray bars). The putative Zn binding domain, the U1, U2, U3, and U4
DISCUSSION
In this report, we describe the cloning of a novel cellulose
synthase required for cell elongation in roots and darkgrown hypocotyls in Arabidopsis. A role for PRC1 (CesA6) in
the synthesis of cellulose was supported by two main ob-
domains, the plant conserved region (PCR), and the HVR are predicted to face the cytosol (Delmer, 1999). The positions of the mutations are indicated. N, N terminus; C, C terminus.
PRC1 Encodes a Cellulose Synthase
2417
tance to strong alkali and the absence of iodine staining also
ruled out the presence of starch in this fraction. The residue
therefore must consist primarily of cellulose. This was further confirmed by labeling experiments, which showed that
less 14C-glucose was incorporated into the nitric/acetic
acid–resistant fraction in prc1 than in the wild type.
A cellulose deficiency was also detected by FTIR microspectroscopy. The advantage of this technique is that
the cell wall composition can be assessed at a microscopic
level in one or a few cells, whereas chemical methods unavoidably provide an average view of many different cell
types. PC analysis of absorption spectra obtained from an
area corresponding exclusively to epidermal and cortical
cells of dark-grown hypocotyls revealed a cellulose deficiency in mutant cell walls. Similar results were obtained
when dark-grown hypocotyls of wild-type seedlings treated
with cellulose synthesis inhibitors such as DCB or isoxaben
were compared with those of untreated controls, confirming
the validity of the method (Fagard, 1999).
PRC1 a Member of the Cellulose Synthase Family
Figure 7. prc1-8 and rsw1-10 Have Similar, and Additive, Hypocotyl
Phenotypes.
Seedlings were grown for 4 days in the dark. Left to right: wild type
(wt), prc1-8, rsw1-10, and double mutant. The prc1-8 and rsw1-10
mutants showed a similar decrease in hypocotyl length compared
with wild-type seedlings, whereas the double mutant showed an
even shorter hypocotyl. Bar ⫽ 2 cm.
servations: prc1 mutants display a cellulose deficiency, and
the PRC1 gene is a member of the cellulose synthase catalytic subunit family.
Prc1 Phenotype a Result of Cellulose Deficiency
Chemical analysis of the wall composition confirmed a cellulose deficiency in mutant dark-grown seedlings. Indeed,
after successive extractions with 0.1 and 4 M KOH, the residual fraction in prc1-1 was reproducibly 25% less than in
wild-type seedlings. This is probably an underestimate, because not all cell types of the dark-grown seedlings were affected by the mutation. This residual fraction was highly
enriched for glucose, which in theory could indicate an
enrichment for (1→3)-␤-glucan (callose), (1→4)-␣-glucan
(starch), or (1→4)-␤-glucan (cellulose). Specific staining with
aniline blue failed to detect a substantial accumulation of
callose in wild-type or mutant hypocotyl cells. The resis-
All six mutant alleles sequenced contained premature stop
codons in CesA6, which in each case removed a major part
or even all of the catalytic domain. The prc1 phenotype is
therefore caused by null alleles of the CesA6 gene. This
gene belongs to family 2 of glycosyl transferase, which also
contains bacterial cellulose synthases. Previous mutant
analysis has shown that at least two other family members,
CesA1 (Arioli et al., 1998) and CesA7 (Taylor et al., 1999),
are required for normal cellulose synthesis in Arabidopsis.
Also, the disappearance of rosettes from scanning electron
microscopy images of freeze-fractured plasma membranes
at the restrictive temperature in rsw1 (mutant for CesA1) and
the immunolabeling of V. angularis rosettes with antibodies
raised against a CesA family member provide strong evidence for a role of these genes in the synthesis of cellulose.
A recent study with virus-induced gene silencing in Nicotiana
benthamiana also showed that the silencing of one or several
members of the CesA family specifically led to cellulose deficiencies (Burton et al., 2000). The final proof will consist of
the in vitro demonstration of cellulose synthesis activity for
products of the CesA gene family. In the absence of an in
vitro assay, the sequence similarity and the cellulose deficiency of prc1 mutants provide strong evidence that CesA6
encodes another cellulose synthase catalytic subunit.
Other Cell Wall Changes in prc1 Mutants
Not only did chemical analysis reveal a deficiency in cellulose, but also mutant walls showed a 50% increase in the
hemicellulose-enriched fraction, the composition of which
did not differ between wild-type and mutant plants. Only a
slight decrease, if any, was observed for the pectin-enriched
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and no additional postembryonic divisions take place, except for those that contribute to the formation of stomatal
guard cells. Because no gapped walls were observed in immature or mature embryos or in light-grown hypocotyls, the
defect must arise during cell expansion. The simplest explanation would be that radial walls, the architecture of which is
not compatible with wall expansion, rupture as a result of
the increased radial expansion caused by the cellulose deficiency. That would be unlikely, however, because several
other radial expansion mutants did not show gapped walls
(Nicol et al., 1998; Bichet et al., 2001). An alternative explanation is that wall expansion, presumably mediated by
Figure 8. No Gene Dosage Effect Is Observed for CesA1 and
CesA6.
Hypocotyl lengths of dark-grown prc1-8 seedlings, mutant for
CesA6, rsw1-10 seedlings, mutant for CesA1, and the progeny of a
plant homozygous for rsw1-10 and heterozygous for prc1-8 were
measured. In the segregating population (SP), a discrete 1:3 segregation of the additive phenotype was observed. No gene dosage effect was observed. Indeed, an intermediate hypocotyl length for half
of the seedlings, those corresponding to prc1-8 heterozygotes,
would be expected if the dosage of the CesA6 gene were critical for
hypocotyl growth.
fraction. Interestingly, in prc1-1, this fraction showed a decreased galactose content without alterations in the content
of uronic acid or other sugars. In primary walls, galactose is
found mainly on galactan side chains of RG1 and on the
side chains of arabinogalactan proteins. However, given the
unaltered arabinose content, the simplest explanation is that
the cellulose deficiency in prc1 caused compensatory
changes in pectin composition, that is, a decrease in the
number or length of the galactan side chains of RG1. We
observed similar changes in cell walls of kor, another cellulose-deficient mutant (I. His, A. Driouch, F. Goubet, and H.
Höfte, unpublished data). The molecular mechanisms underlying these secondary changes and their functional significance, if any, remain to be determined. Finally, the
chemical analysis did not provide evidence for the accumulation of substantial amounts of alkali-extractable nonhemicellulosic glucan in prc1, whereas such accumulation was
reported for rsw1.
An intriguing observation was the presence of gapped
walls in endodermal, cortical, and epidermal cells of prc1
hypocotyls and roots. This is unlikely to be an artifact of the
sample preparation, because the same defect was observed when using three different sample preparation procedures, including scanning electron microscopy, which
does not involve tissue fixation. This defect is reminiscent of
the stubs observed in cytokinesis mutants. Cortical and epidermal hypocotyl cells essentially have an embryonic origin,
Figure 9. Members of the CesA Family Exhibit Functional Specialization.
Shown is a phylogenetic tree based on the HVR (see text and Figure
5) of protein sequences of Arabidopsis and cotton CesA family
members. Alignment data are from bootstrap values sampled 100
times and used to construct the consensus tree shown. Numbers
are bootstrap values and indicate the number of trees in which the
proteins to the right of the bootstrap values clustered together. Gh
EST-AI727450 was included as an outlier. The five other cotton
genes did not cluster together but instead clustered with one or two
Arabidopsis sequences, suggesting that the CesA amino acid sequences diverged before the divergence between Brassicaceae and
Malvaceae; that, in turn, suggests a functional specialization of most
CesA isoforms.
At, Arabidopsis thaliana; Gh, Gossypium hirsutum.
PRC1 Encodes a Cellulose Synthase
2419
wall-loosening proteins such as expansins, continues in the
absence of sufficient cellulose synthesis to maintain a constant wall thickness. This eventually may lead to the rupture
of the walls. Interestingly, the space between the wall stubs
often contained wall material, composed of at least xyloglucan and pectin, as found with specific antibodies (data not
shown). These results suggest that at least in mutant walls,
impaired cellulose synthesis does not feed back to the wallloosening machinery and the synthesis of other wall polysaccharides.
Functional Specialization within the Cellulose Synthase
Gene Family
Large gene families are more the rule than the exception in
plants, including Arabidopsis. Reverse genetics frequently
has failed to associate clear-cut mutant phenotypes with
mutations in individual genes within such families (Gilliland
et al., 1998; D. Bouchez, personal communication). This
suggests that many gene family members may have at least
partially redundant roles. The CesA gene family is atypical in
this respect, in that strong mutant phenotypes have been
observed for at least three genes. Tissue-specific expression may explain specific phenotypes in some cases, for example, RSW1 in cells producing primary walls and IRX3 in
xylem elements producing secondary walls. However, the
conservation of specific amino acid sequence motifs, such
as the HVR, between species suggests a functional specialization of different isoforms. We think it likely that the HVR
domain interacts with other proteins with specific roles in
different cell types or environmental conditions.
Another complication involves the observation that in
some cell types at least two isoforms are required for normal
cellulose synthesis. Indeed, rsw1-1 is affected in all cell
types at the restrictive temperature, including in cells in the
hypocotyl and root (Arioli et al., 1998). Cell expansion defects in all expanding cell types, including those in the central cylinder, were also observed in roots and hypocotyls of
rsw1-10. The phenotype of rsw1-10 is less severe than
rsw1-1 at the restrictive temperature (data not shown),
which suggests that rsw1-10 is a leaky allele. Null alleles of
PRC1 also showed expansion defects in endodermis, cortex, and epidermis of dark-grown hypocotyls and roots, indicating a requirement for both gene products to support
normal cellulose synthesis and cell expansion in these cell
types. The simplest explanation for this observation is a
quantitative one: to sustain the demand for cellulose in the
cell, a sufficiently large pool of CesA protein is required,
which in turn requires synthesis from both genes. The
strictly recessive phenotypes for both genes do not support
a strong dependence on dose. An even stronger argument
against this hypothesis is the absence of a gene dosage effect in a population segregating for the prc1-8 mutation in a
genetic background homozygous for rsw1-10. The results
suggest a strict requirement for two distinct isoforms rather
Figure 10. PRC1 and RSW1 Have Similar Transcript Profiles.
Total RNA of different samples was hybridized with labeled probes
corresponding to a variable sequence at the N-terminal end of CesA
proteins.
(A) PRC1 probe.
(B) RSW1 probe.
(C) Ethidium bromide–stained agarose gel of the same samples.
Lane 1, 1-day-old germinating seeds; lane 2, 4-day-old dark-grown
seedlings; lane 3, 4-day-old light-grown seedlings; lane 4, elongating inflorescence stem sections; lane 5, mature leaves; lane 6, roots;
lane 7, mature flowers.
than a certain pool size of CesA protein. This implies that
different CesA isoforms need to be present at different times
in the same cell or at different locations within the cell. Alternatively, CesA isoforms may act in concert or might form
heterodimers. Immunolocalization studies with specific antibodies or with epitope-tagged versions of RSW1 (CesA1)
and PRC1 (CesA6) should clarify this issue.
A final intriguing observation is the light-dependent conditional phenotype in mutant prc1 hypocotyls. We previously
showed that the rescue of the hypocotyl phenotype is mediated by phytochrome (Desnos et al., 1996). The simplest explanation for this phenomenon is that phytochrome controls
the expression of one or more other CesA isoforms, which
either replace PRC1 (CesA6) in light-grown hypocotyls or
have a function redundant with it. These observations indicate that phytochrome acts on the cellulose synthesis machinery and suggest a connection between phytochrome
signaling and cell wall–related processes involved in growth
control. RNA gel blot experiments thus far do not provide
evidence for the regulation of PRC1 mRNA quantities by
light. We are currently investigating the control by phytochrome of the expression of other CesA family members.
METHODS
Plant Material and Growth Conditions
Wild-type Arabidopsis thaliana plants were of the Columbia (Col-0),
Wassilewskija (Ws), or Landsberg erecta (Ler) ecotype. Alleles prc1-1
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The Plant Cell
through prc1-4 have been described previously (Desnos et al., 1996)
and were of the Col-0 ecotype; these lines were all backcrossed four
times. Alleles prc1-5 through prc1-7, prc1- 9, and prc1-11 (T. Desnos,
F. Nicol, and H. Höfte, unpublished data; P. Benfey and M.-T.
Hauser, personal communication) were of the Col-0 ecotype; these
lines were backcrossed two, two, one, two, and one time, respectively.
Mutant rsw1-1 was of the Col-0 ecotype (Arioli et al., 1998). Mutants
rsw1-10, prc1-8, prc1-10, prc1-12, and prc1-19 were isolated from
T-DNA–mutagenized populations and were of the Ws ecotype.
For growth in vitro, plants were grown on Petri dishes containing
Murashige and Skoog (1962) medium (Estelle and Somerville, 1987)
without sucrose at 25⬚C. For dark growth, seeds were cold-treated
for 48 hr and then exposed to fluorescent white light (200
␮M.m⫺2.sec⫺1) for 2 hr to synchronize germination, and finally the
plates were wrapped in three layers of aluminum foil. For culture in
the light, seeds were cold-treated for 48 hr and then placed under
fluorescent white light (200 ␮M.m⫺2.sec⫺1) in 16-hr-light/8-hr-dark cycles. The time of measurement or collection of tissue was counted as
days after the beginning of the light treatment. For cellulose incorporation assays, 200 Arabidopsis seedlings were grown at 20⬚C in the
dark for 3 days in 10 mL of liquid medium containing 0.5% glucose.
Seeds were cold-treated for 48 hr and then exposed to fluorescent
white light (200 ␮M.m⫺2.sec⫺1) for 5 hr to synchronize germination,
after which the flasks were wrapped in three layers of aluminum foil.
samples were dried at 37⬚C for 20 min. Two spectra of each seedling
were collected on a Bio-Rad FTS-40 FT spectrometer in the middle region of the hypocotyl, avoiding the central cylinder, in a 60- ⫻ 40-␮m
window. Because absorbance varies with sample thickness, all data
sets were corrected for baseline and normalized for area before statistical methods were applied. Exploratory principal components analysis was performed with Win-Discrim software (E.K. Kemsley, Institute
of Food Research, Norwich, UK). Reference infrared absorption spectra of cellulose and other ␤-glucans were obtained from the literature
(Tsuboi, 1957; Liang and Marchessault, 1959; Séné et al., 1994).
Preparation of Cell Wall Material
The plants were incubated for 30 min in 90% ethanol at 65⬚C to inactivate enzymes. The tissues were then ground in a Tenbroeck glass
Potter–Elvehjem homogenizer (Janke and Kunkel IKA Labortechnik,
Staufen, Germany). The homogenate was centrifuged at 2300g for
15 min. The pellet was washed with ethanol (three or four times),
methanol/chloroform (2:3 [v/v], overnight), and ethanol again. The remaining pellet, which contained the cell wall, was dried overnight at
80⬚C. The yield of cell walls was defined as the weight (in grams) of
dry cell walls per 100 g of fresh plants.
Extraction of Polymers
Measurement of Hypocotyl and Root Lengths
The growth of the seedlings was arrested by adding an aqueous solution of 0.4% formaldehyde. Hypocotyls and roots were spread on agar
plates, and their image was captured with a digital camera. Lengths
were measured by using image analysis software (Optimas 5.2; IMASYS, Surennes, France), as described by Gendreau et al. (1997).
Pectic and hemicellulosic polymers were solubilized successively
from cell walls by treatments with 0.1 M KOH at 20⬚C (three times for
1 hr) and 4 M KOH at 20⬚C (three times for 1 hr), respectively. The remaining pellet was washed with water until the pH of the supernatant
was equal to that of water. The KOH extracts were neutralized with
HCl, dialyzed against water with a molecular weight cutoff of 1000,
and lyophilized.
Cross-Sections
Sugar Composition
For light microscopy, seedlings were fixed in PBS buffer containing
4% paraformaldehyde and 0.2% glutaraldehyde and were embedded in historesin (Technovit 7100; Kulzer, Wehrheim, Germany), according to the manufacturer’s instructions. Sections 3 ␮m thick were
cut by using a Jung RM2055 microtome (Leica Instruments, Nussloch, Germany). The cross-sections were stained with a 0.005%
aqueous solution of Calcofluor (fluorescent brightener 28; Sigma) for
2 min and visualized under UV light with a Microphot FXA microscope (Nikon, Champigny-sur-Marne, France).
For transmission electron microscopy, seedlings were fixed with a
2.5% glutaraldehyde solution in phosphate buffer and then stained
with a 2% osmium tetroxide solution. Seedlings were then embedded
in Epon resin (TAAB 812, TAAB Laboratories Equipment Ltd., Reading, UK). Ultrathin sections (100 nm thick) were cut with a Reichert
(Leica Instruments) Ultracut E ultramicrotome and deposited on 300mesh copper–palladium grids. Samples were stained a second time
with a 5% uranyl acetate solution followed by a 0.5% lead citrate solution and visualized with an EM 420 transmission electron microscope (Philips, Eindhoven, The Netherlands) at 80-kV acceleration.
Fourier Transform Infrared Spectroscopy
The seedlings (10 for each condition) were pressed onto a barium fluoride window and rinsed with water to remove cytoplasmic debris. The
Neutral sugar composition was determined as described by Englyst
and Cummings (1984). Inositol was added to samples as an internal
reference. Uronic acid contents were determined as described by
Thibault (1979).
Cellulose Incorporation Assays
Two hundred Arabidopsis seedlings, grown in liquid culture in the
dark, as described above, were washed three times with 15 mL of
glucose-free growth medium before being resuspended in 1 mL of
growth media containing 14C-glucose (NEN Research, Boston, MA),
1.0 ␮Ci.mL⫺1 . The seedlings were then incubated for 1 hr in the dark
at 15⬚C in glass tubes. After treatment, the seedlings were washed
three times with 6 mL of glucose-free growth medium and then extracted with 5 mL of boiling absolute ethanol for 20 min. This step
was repeated three times. Next, seedlings were resuspended in 3 mL
of chloroform/methanol (1:1 [v/v]), extracted for 20 min at 45⬚C, and
finally resuspended in 3 mL of acetone for 15 min at room temperature with gentle shaking. The remaining material was resuspended in
500 ␮L of an acetic acid/nitric acid/water solution (8:1:2 [v/v/v]), as
described by Updegraff (1969), for 1 hr in a boiling water bath. Acidsoluble material and acid-insoluble material were separated by vacuum filtration through glass microfiber filters (GF/A; 2.5 cm diameter;
PRC1 Encodes a Cellulose Synthase
Whatman, Maidstone, UK), after which the filters were washed with
1.5 mL of water. The acid solution and water wash constitute the
acid-soluble fraction. Finally, the filters were washed with 30 mL of
water and 20 mL of ethanol, yielding the acid-insoluble fraction,
which is essentially crystalline cellulose. The amount of label in each
fraction was determined by scintillation counting in a Betamatic scintillation counter (Kontron Instruments, Montigny le Bretonneux,
France) using Ultima-Flo AP (Packard, Groningen, The Netherlands)
as the scintillation liquid. The percentage of label incorporation was
expressed as 100 ⫻ the ratio between the amount of label in the cellulosic fraction and the amount in the acid-soluble plus cellulosic
fractions.
Genetic Analysis and Map-Based Cloning
The prc1-1 mutant (ecotype Col-0; backcrossed three times to Col-0)
was crossed to line KanR5 (ecotype C24), which bears a T-DNA
(conferring resistance to kanamycin) inserted close to marker LFY3
(Van Lijsebettens et al., 1996) at the bottom of chromosome 5 (Lister
and Dean, 1993). F2 seedlings were sown on Murashige and Skoog
medium supplemented with 50 mM kanamycin and grown for 3 days
in total darkness. Of these, 1632 seedlings with short hypocotyls
were scored and cultured for 3 more days in white light. Individuals
with the prc1 phenotype and resistant to kanamycin were considered
recombinant between the prc1-1 mutation and the T-DNA insertion.
These plants were transferred to the greenhouse, and genomic DNA
was extracted from leaves and flower buds, as described previously
(Bouchez and Camilleri, 1998).
To determine the position of the prc1-1 mutation, DNA of the recombinant plants was analyzed with restriction fragment length polymorphism marker 5H1-L; this marker was designed by isolating the
left border of yeast artificial chromosome 5H1 using the vectorette
technique (Riley et al., 1990) and looking for polymorphism between
the Col-0 and C24 ecotypes by DNA gel blotting with different restriction enzymes. The MUB3.12 and MUB3.p2B cleaved amplified
polymorphic sequence (CAPS) markers were designed by polymerase chain reaction amplification of introns from genes 12 and
p2B of P1 clone MUB3 (GenBank accession number AB010076) and
digestion of the polymerase chain reaction products, amplified from
genomic DNA of Col-0 and C24 ecotypes, with pools of restriction
enzymes (40 were tested in all). The MUB3.12 marker (specific oligonucleotides: forward, CGCCGACGGAGAAATGACCAAG; reverse,
TCACCAACACGAATGCGAGTACGA) is polymorphic for an EcoRI
restriction site that is present in ecotype Col-0 but absent in C24 and
Ler, whereas the MUB3.p2B marker (specific oligonucleotides: forward, CTCTCCCAAAGTAATCGTGCTA; reverse, TACCAGATGAACAGGGCTAAGT) is polymorphic for an AccI site that is present in C24
but not in Col-0.
The prc1-1 mutant was also crossed to a wild-type plant of the Ler
ecotype. F2 seedlings were grown in the dark on Murashige and
Skoog medium, and individuals homozygous for the prc1-1 mutation
were selected on the basis of their short hypocotyls. These plants
were transferred to the greenhouse, and genomic DNA was extracted from leaves and flower buds (Edwards et al., 1991). Recombinant individuals were identified by using CAPS markers based on
partial sequences of markers ATPK41B (Recombinant inbred
marker; Lister and Dean, 1993) and ATHB-5 (GenBank accession
number X67033). Specific oligonucleotides (forward, TGTGATCAGAAATGGATCG; reverse, CTTCCTGTGTTGCTCTCGTT) amplified an
ⵑ700-bp fragment at the ATPK41B locus, which was found, by test-
2421
ing with 40 commonly used restriction enzymes, to be cleaved by
HinfI in the Ler allele but not in the Col-0 allele. For optimal resolution
of the polymorphism, the polymerase chain reaction fragments were
also cut with the restriction enzyme DraI. Other specific oligonucleotides (forward, TAACCAGCTCATCAACCAAAC; reverse, AAGATGGAAACAGAGAATTGACTA) amplified an ⵑ850-bp fragment at the
ATHB-5 locus, which is cleaved by the restriction enzyme AvaI in the
Col-0 allele but not in the Ler allele. Recombinant inbred marker Annexin (ve028; Lister and Dean, 1993) was converted to a CAPS
marker (specific oligonucleotides: forward, TCCACCAAGATGTTCACCAAGA; reverse, CGTTCCATGTCAACTTCAGTTC) by using the
restriction enzyme TfiI. Finally, a restriction fragment length polymorphism marker (2F2-E12) was found on the proximal part of P1 clone
MXK3, which is polymorphic for an HpaII restriction site between Ler
and Col-0.
RNA Gel Blotting
RNAs were extracted as described previously (Brusslan and Tobin,
1992). RNA gel blotting on Hybond N membranes and labeling of the
probes by random priming in the presence of 50 ␮Ci of 32P-dCTP was
performed according to the manufacturer’s instructions (Amersham,
Little Chalfont, UK). The membrane was hybridized overnight at 42⬚C
in 10 mL of SDS-rich buffer, as described by Church and Gilbert
(1984) with 50% formamide and 100 ␮g/mL salmon sperm DNA after
overnight prehybridization. Washes were performed at 65⬚C in 2 ⫻
SSC (1 ⫻ SSC is 0.15 M NaCl and 0.015 M sodium citrate) plus 0.1%
SDS for 15 min and again for 15 min in 1 ⫻ SSC plus 0.1% SDS
(once for the RSW1 probe and twice for the PRC1 probe).
The RSW1 probe was prepared with a 300-bp-long EcoRV-HincII
fragment covering the N-terminal variable region, which is specific
for each CesA (positions 322 to 622 from ATG). The PRC1 probe was
prepared with a 379-bp-long BglII-HindIII fragment covering the
same region (positions 237 to 616 from ATG). The specificity of the
probes was tested on dot blots of cloned cDNAs corresponding to
Arabidopsis cellulose synthase genes, in the same hybridization conditions as those used for the RNA gel blots. The hybridization signal
of the RSW1 probe was at least 10-fold greater for RSW1 than for
PRC1. The hybridization signal of the PRC1 probe was at least 10fold greater for PRC1 than for the very similar sequence CesA5 and
at least 100-fold greater than for RSW1 (data not shown).
Phylogenetic Analysis
Unrooted trees were built by using an alignment of a variable region
corresponding to amino acids 690 and 743 in PRC1 obtained with
the program ClustalW (http://www.infobiogen.fr/services/analyseq/
cgi-bin/clustalw_in.pl). This alignment was used by the algorithm
PROTPARS (also available at the above-mentioned website) to construct a consensus tree from 100 bootstrapped data sets.
ACKNOWLEDGMENTS
We thank Brigitte Gelie, Jocelyne Kronenberger, Joël Talbotec,
Sigolène Müller, Audrey Hantiu, and Olivier Grandjean for technical
assistance; Estelle Aletti for providing the data described in Figure 7;
Hervé Vaucheret for enthusiastic discussions and critical reading of
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The Plant Cell
the manuscript; Phillip Benfey and Marie-Therese Hauser for providing prc alleles; and Reginald Wilson for insightful advice regarding
FTIR. We thank Vincent Chiang for kindly providing the AraxCelA
cDNA clone. Jean-François Thibault and Marc Lahaye are thanked
for thoughtful advice concerning sugar chemistry and for making
their equipment available. The Arabidopsis Biological Resources
Center and the Nottingham Stock Centre are thanked for providing
seed and DNA stocks. This work was financed in part by Framework
4 Grant PHOTARCH (No. BIO4-CT97-2124) from the European Economic Community; Grant No. A.I.P. 00176 from the Groupement de
Recherche, Génomes et Mutants d’Arabidopsis; and Grant No. ACCSV 9501006 from the Ministère de la Recherche et de la Technologie.
T. Desnos and M.F. were bursaries from the Ministère de la Recherche
et de la Technologie.
Desnos, T., Orbovic, V., Bellini, C., Kronenberger, J., Caboche,
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PROCUSTE1 Encodes a Cellulose Synthase Required for Normal Cell Elongation Specifically in
Roots and Dark-Grown Hypocotyls of Arabidopsis
Mathilde Fagard, Thierry Desnos, Thierry Desprez, Florence Goubet, Guislaine Refregier, Gregory
Mouille, Maureen McCann, Catherine Rayon, Samantha Vernhettes and Herman Höfte
PLANT CELL 2000;12;2409-2424
DOI: 10.1105/tpc.12.12.2409
This information is current as of September 3, 2009
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